Western Blotting Protocol

Buffers:

TBS:  25 ml of 1 M Tris-7.5, 30 ml of 5 M NaCl, bring volume up to 1000 ml with ddwater

TBS-T:  TBS + 0.5 ml of Tween 20

5X SDS-PAGE running buffer:  15.1 g Tris base, 94 g glycine in 900 ml water, then add 50 ml of 10% solution of SDS, and adjust volume to 1000 ml with ddwater

Blocking solution:  5 g nonfat dry milk in 100 ml of PBS

Transfer Buffer:  5.82 g Tris base + 2.93 g glycine + 0.375 g SDS (or 3.75 ml of a 10% solution of SDS) + 200 ml methanol, then bring volume up to 1 liter with ddwater

Prepare cell extracts using an appropriate protocol.

1. Measure total protein in extract by the Bradford method.  Plan on loading approximately 5 μg of protein per lane on the polyacrylamide gel.

2. Add an equal volume of SDS sample buffer and boil the sample on a 100℃ block for 5 min.

3. Apply the sample in about 10 μl volume to an appropriate percentage (typically 4-20% gradient) SDS-PAGE mini gel, along with visible protein marker (5 ul) and appropriate positive and negative controls.  Run gel using 1X SDS running buffer.

4. Run gel at 25 mA (constant) for 30 min to 1 hr. (or until blue dye just runs off of the gel)

5. Transfer onto immobilon P membrane using the SD semi-dry transfer apparatus, per apparatus instructions.

6. After transfer, ensure that all the protein has transferred, as assessed by transfer of the visible protein marker onto the immobilon P membrane.

7. Block in 75 ml of blocking solution overnight at 4 ℃(alternatively, block at room temp for 1 h on a shaking platform).

8. Wash 3 times with TBS-T, each for 10 min.

9.Make dilution of primary antibody (usually 2 μl in 10 ml of TBS), and add to membrane, shake on a platform for 1 h at room temp. 

Make sure that membrane is entirely covered with antibody solution.

10.  Discard antibody solution and wash with TBS-T 3 times for 10 min each.

11.  Make dilution of secondary antibody (usually 1 μl in 10 ml), and cover membrane with antibody solution and shake on a platform for 1 h at room temp.

12.  Discard antibody solution, and wash membrane with TBS-T 4 times for 10 min. each.

13.  Detection using the ECL/ECLplus kit.

DUAL ELISPOT PROTOCOL

staining provides the opportunity to detect two target proteins at the same time in the ELISPOT cell assay. An advantage of this is that it enables the researcher to detect a marker protein to determine which cell type is secreting the protein, as well as the secreted protein of interest.
The following dual ELISPOT procedure describes a protocol using an FITC-conjugated primary antibody and a biotinylated primary antibody. Those are in turn recognized by anti-FITC HRP and streptavidin-AP conjugates. PVDF-bottomed-well plates are then incubated first with AEC substrate buffer, washed and subsequently incubated with BCIP/NBT. The colored spots will either be red/brownish blue/purple so the two target proteins can be distinguished.

Materials and Reagents:
1.Detection antibody
2.Streptavidin alkaline phosphatase conjugate
3. Phosphate buffered saline (10 x Concentrate solution).
For 1 liter weight : 80g NaCl ; 2g KH2PO4; 14.4g Na2HPO42H2O. Add distilled water to 1 liter. Check that pH is 7.4+/- 0.1. Dilute the solution to 1X before use.
4. 2% dry skimmed milk in PBS
For one plate dissolve 0.2 g of powder in 10 mL of 1X diluted PBS.
5. 1% BSA in PBS
For one plate dissolve 0.2 g of BSA in 20 mL of 1X diluted PBS.
6. 0.1% Tween in PBS
For one plate dissolve 100􀀁l of Tween 20 in 100 ml of 1X diluted PBS.
7. 70% ethanol in water
For one plate mix 7 ml of ethanol with 3 ml of distilled water.
8. AEC Buffer
For one plate mix 1 ml of ACE Buffer A with 9ml of distilled water. Then add 200􀀁l of ACE Buffer B.
4. ELISPOT 96 well plates
(provided with Abcam ELISPOT kits)

AFLP Protocol

Restriction digestion
Master mix preparation:
Prepare a master mix of the following per sample, plus 5 to 10% extra to allow for pipetting loss.
5X R/L buffer (see page 4 for recipe) 6.0 μl
EcoR I (12 U @20 U/μl) 0.6 μl
Mse I (8 U @ 4 U/μl) 2.0 μl
Water to 10 μl
Reaction time:
Incubate for at least 1 hr. at 37°C, but not longer than 3 hrs, before proceeding with the ligation.

Adapter ligation
Master mix preparation
If 10 μl of the restriction reaction was removed for gel analysis, prepare a ligation master mix of the following, per sample, plus enough for 5-10 extra samples:
EcoRI adapter (@ 5 pMol/μl) 0.5 μl
MseI adapter (@ 50 pMol/μl) 0.5 μl
ATP (10 mM, pH 8.0) 0.5 μl
5X R/L buffer 1.0 μl
sdH2O 2.0 μl
T4 DNA ligase (0.5 Weiss U @ 1 U/μl) 0.5 μl
TOTAL 5.0 μl
Reaction time:
Incubate for at least 3 hrs. (preferably overnight) @ 37° C. After incubation, dilute each R/L mix 1:10 with sdH₂O.

Preamplification
Master mix preparation
Prepare a master mix with the following amounts per sample, plus 5-10 samples extra:
10X PCR buffer 3.0 μl
dNTP mixture (2.5 mM ea.) 2.4 μl
E primer (@ 50 ng/μl ≅ 8.3μM) 1.0 μl
M primer (@ 50 ng/μl ≅ 8.3μM) 1.0 μl
Taq polymerase (@ 5 U/μl) 0.4 μl
sdH₂O 19.2 μl
μl per sample 27.0 μl

Important: If pre-amplifications are to be done in 96-well PVC plates instead of polycarbonate or polypropylene plates or tubes, it may be necessary to add 2.0 μl of 10 μg/μl BSA per sample to the Taq-buffer mix, and decrease the amount of H2O to 5.76 μl per sample. DO NOT use a hot start PCR protocol, or a hot start polymerase (e.g. AmpliTaq Gold), for the pre-amplification! The adapters are non-phosphorylated, so only the top strand is ligated. The bottom strand of the adapter will separate from the rest of the template first and must be re-synthesized during the initial heating stage. This requires polymerase activity during the initial heating.
PCR program
Place reactions in the thermocycler, and run the following PCR amplification profile:
28 cycles: 15 sec @ 94°C denaturation
30 sec @ 60°C annealing
60 sec + 1sec/cycle @ 72°C extension
1 cycle: 2 min @ 72°C final extension
hold: 4°C

Final AmplificationMaster mix preparation (single dye reactions)
For each E/M primer combination to be used, prepare the following master mix, per sample, in an Eppendorf tube, plus enough for 5-10 samples extra:
10X PCR buffer 2.0 μl
dNTP mixture (2.5 mM ea) 1.6 μl
IRD-labeled E-primer (@ 6 ng/μl ≅ 1μM) 0.83 μl
M-primer (@ 50 ng/μl ≅ 8.3μM) 0.6 μl
Taq polymerase (@ 5U/μl) 0.24 μl
dH₂O 9.73 μl
Final volume 15.0 μl

Cleaved Amplified Polymorphic Sequences (CAPS)

Cleaved Amplified Polymorphic Sequences (CAPS) polymorphisms are differences in restriction fragment lengths caused by SNPs or INDELs that create or abolish restriction endonucleaserecognition sites in PCR amplicons produced by locus-specific oligonucleotide primers.

How It Works

The CAPS assay uses amplified DNA fragments that are digested with a restriction endonuclease to display RFLP.

Unique sequence primers are used to amplify a mapped DNA sequence from two related individuals (for example, from two different inbred ecotypes), A/A and B/B, and from the heterozygote A/B. The amplified fragments from A/A and B/B contain two and three RE recognition sites, respectively. In the case of the heterozygote A/B, two different PCR products will be obtained, one which is cleaved three times and one which is cleaved twice. When fractionated by agarose or acrylamide gel electrophoresis, the PCR products digested by the RE will give readily distinguishable patterns. Some bands will appear as doublets.

Advantages

• Most CAPS markers are co-dominant and locus-specific.
• Most CAPS genotypes are easily scored and interpreted.
• CAPS markers are easily shared between laboratories.
• CAPS assay does not require the use of radioactive isotopes, and it is more amenable, therefore, to analyses in clinical settings.

Developing CAPS markers

• Sequence the RFLP probe.
• Design primers to amplify 800–2,000-bp DNA fragments. Targeting introns or 3' untranslated regions should increase the chance of finding polymorphisms
• The PCR product is cloned and sequenced.
• PCR amplify DNA fragments from target genotypes, separately digest the amplicons with one or more restriction emzymes.
• Screen the digested amplicons for polymorphism on gels stained with ethidium bromide.

QRT-PCR protocol

General notes:

       Consistency is crucial to the accuracy of QRT-PCR, so the extra steps in this protocol are necessary. In general, be extra careful to treat every sample the same. Always use filter tips, and do no aspirate when pipetting (only go down to the first stop). Avoid bubble formation, which can accelerate the degradation of the cDNA. The SYBR Green is photosensitive, so the mixing is done in the “dark” room in Judy Manning’s lab, which is also the location of the iCycler machine. For the final spin of the 96-well plate, wrap the plate in foil before bringing it into the lab, and minimize exposure to light. Samples are done in triplicate to ensure that the technique is good.

Before beginning protocol

1. Determine what templates will be used, and which will be used in the dilution series. At least one template should have a dilution series of five 10-fold dilutions.
2. For the primer set being used, determine the best annealing temperature by doing end-point PCR using a gradient for the annealing temperature. Pick the highest temperature with the best band. Use this temperature for the QRT-PCR reaction. Because of this, different primer sets must be done in separate iCycler runs, unless the best annealing temperature is the same for two sets.
3. Sign up for the iCycler ahead of time. The sign-up sheet is currently at the end of the McFall-Ngai lab bench on the left side of the aisle, as you face the windows.
4. It is a good idea to get the iCycler program and well designations done before starting work with the samples.

Protocol:

1. Make master mix: Multiply the volumes per reaction by the number of reactions, and add 10%.

       Component                                Volume added per reaction

       2x iQ SYBR green                        12.5 ul

       Forward Primer (10 uM)               0.5 ul   

       Reverse Primer (10 uM)                   0.5 ul

       Sterile Distilled H₂O                            10.5 ul

      

       Total volume                               24.0 ul

Note: the amount of primers used will vary with the stock concentration. This assumes that 1 ul template is added per reaction. If this is not the case, adjust the volumes accordingly.

2. Vortex mix briefly, then do a short spin to remove any bubbles.

3. Prepare PCR tubes with 3 ul of each template.

5. Add 72 ul of master mix to each tube.

6. Vortex and spin tubes briefly.

7. Place 23 ul of each sample in the appropriate wells in a 96-well plate (less than the full amount is used to make sure that there is enough for all three wells, and to avoid bubbles). Make sure to put the samples in the correct well, as only those wells designated by the program will be read and recorded.

8. Carefully put clear cover on the plate. Use paddle to smooth down – do not touch cover.

9. Wrap plate in foil. Find similarly filled plate (there’s probably a pile) for a balance. Take into main lab, and spin down in Sorval Legend RT at *** for **** minutes.

10.  Return to iCycler room. Keep plate in cold block. Begin iCycler run, and when the block gets up to temperature, pause the program and place the plate in the block. Well A1 goes in the top left as you are facing the machine. Unpause the program, and allow to run. Do not turn on the lights in the room until the run is over.

Recovering Virus Stocks from Frozen Cell Pellets

If your recombinant baculovirus stocks have been lost or have "gone off", virus can usually be recovered from frozen pellets of baculovirus-infected insect cells. Virus in these pellets is stable for many years, especially if stored at -70 C or less.

Procedure:

1. Resuspend the frozen pellet in 10 volumes of insect cell growth medium supplemented with 5% FBS. Vortex briefly.
2. Centrifuge at low speed to remove cell debris (1-2000 x g for 5 min.).
3. Filter supernate with 0.45 micron sterile filter (not O.2 micron).
4. Infect insect cells by adding a volume of filtered virus to the cell cultures that is equivalent to 5% of the total media volume. Normally this is done using stationary cultures at about 5E5 cells/ml in a vessel such as a T-125 flask (10-12 ml total per flask) with media containing 2-5% FBS (FBS enhances virus stability).
5. Harvest virus stocks when all cells appear infected, or when they become confluent (3-5 days). Some toxicity, affecting 10-40% of the cells, usually occurs within 24 hours of infection. Re-passage the virus stock as necessary.

Purification of Monoclonal Antibody from Hybridoma Culture Supernatant

Step1: Hyridoma cell culture

• Thaw freezing cell line 9E10 which express secreted monoclonal antibody(subtype IgG1) against Myc epitope in RPMI1640 medium containing 10% FBS and appropriate amount of ampicillin and streptomycin, incubate cell at 37degree with 5% CO2.
• Once cell get into log phase growth, cell should be passaged by every one day, the cell density start at 2x 105/ml. Note: 9E10 and 12CA5 cell are half adhesive , once they are completely confluent, they will immediately start cell death.
• Centrifuge cultured medium at 1000 rpm for 5 minitures, collect supernatant into sterile containers, if necessary add sodium azide up to 0.2%. for several month storage, keep supernatant at 4 degree, otherwise, freeze them at -20 degree.
• Freeze log phase cells for stock, put at least 2 million cell in 1 ml RPMI1640 medium containing 20% FBS,15% DMSO, keep freezing vials at -70 for no more than one month, then transfer them to nitrogen tank.

Step2: Affinity purification of antibody

• Affinity column choosing: protein G for mouse IgG1 and protein A for mouse IgG2a, IgG2b, IgG3
• set appropriate volume of protein G sepharose column according to the common rule: culture supernatant contain 20-50ug/ml antibody, 1ml of wet beads bind approximately 10-20mg antibody, wash column with 100mM Tris-HCL pH 8.0.
• Adjust cell culture supernatant pH by adding 1/10 volume of 1.0M tris-HCL pH8.0, pass it through protein G column at speed of 2ml/min.
• wash column with at least 10 column volume of 100mM Tris-HCL then wash with 10mM Tris-HCL.
• Elute the column with 50 mM glycine(pH3.0), add this buffer stepwise at 1ml per time, collect elute fraction into 1.5 ml eppendorf tube containing 100ul 1M Tris-HCL for immediate neutralization of antibody solution
• During collecting elute fraction, use bradford solution to monitor eluted protein
• Running 5 ul of each fraction on 11% SDS-PAGE gel to check the purity of antibody
• Combine the fractions containing antibody, determine the antibody concentration by measuring OD280nm(1 OD= 0.75 mg/ml)
• Make series dilution of antibody such as 1:500, 1:1,000, 1:2,000, 1:3,000, 1:4,000, perform western blot to determine the optimal titre of the antibody, when doing titration, the antigen should be Myc (for 9E10) or HA (for 12CA5) tagged protein which was confirmed by other Western bolt.